Several approaches have been proposed for improving the biocompatibility of biomaterials useful in medical applications. For example, modifying the biomaterial surface to provide either low polarity or ionic charge or coating the surface with biological substances such as cells, proteins, enzymes, etc., has been used to prevent undesirable protein adhesion. Another approach involves the incorporation of an anticoagulant into the biomaterial, rendering the biomaterial antithrombogenic. A further approach involves the incorporation of various phospholipids into the biomaterial. An additional approach involves the binding of a heparin-quaternary amine complex, or other antithrombotic agent, to the biomaterial surface However, many of these methods have the disadvantage of being nonpermanent systems in that the surface coating is eventually stripped off or leached away. For example, heparin, which is complexed to the biomaterial surface, is not only gradually lost from the polymer surface into the surrounding medium but also has limited bioactivity due to catabolism and its inherent instability under physiological conditions.
Membranes, as self-organizing noncovalent aggregates, offer a model for molecular engineering in which the constituent members can be controlled, modified, precisely defined, and easily assembled. During the past decade, phospholipids differing in chemical composition, saturation, and size have been utilized as building blocks in the design of a variety of structures of complex geometry. Lipid-based cylinders, cubes, and spheres have found applications in both drug delivery and as templates for composite molecularly engineered structures. Surface-coupled bilayers for biosensor applications have also been produced by assembling a layer of closely packed hydrocarbon chains onto an underlying substrate followed by exposure to either a dilute solution of emulsified lipids or unilamellar lipid vesicles (Spinke et al. [1992] Biophys. J. 63:1667-1671; Seifert et al. [1993] Biophys. J. 64:384-393; and Florin et al. [1993] Biophys. J. 64:375-383). In addition, Langmuir-Blodgett techniques have been used as an alternate strategy to construct supported bilayers via a process of controlled dipping of a substrate through an organic amphiphilic monolayer (Ulman, A. [1991] “An Introduction to Ultrathin Organic Films from Langmuir-Blodgett to Self-assembly,” New York, Academic Press). Remarkably, these noncovalent molecular assemblies exhibit a high degree of stability. A force of 26 kT is required to remove a double chained C-16 phosphatidylcholine molecule from a bilayer into water (Cecv, G. and Marsh, D. [1987] “Phospholipid Bilayers,” New York, Wiley; and Helm et al. [1991] Proc. Natl. Acad. Sci. USA 88:8169-8173). This nearly approximates the biotin-streptavidin bond energy of 35 kT and is several orders of magnitude greater than the strength of typical monoclonal antibody-antigen interactions. Thus, the significance of the methodologies of the present invention lies in the ability to engineer relatively robust materials with an unparalleled level of reproducibility and molecular control over surface order and chemistry.
In order to create robust surface structures, most membrane-mimetic systems for blood-contacting applications have been designed as copolymers containing the phosphorylcholine functional group in either side chains or, less frequently, the polymer backbone (Kojima et al. [1991] Biomaterials 12:121; Ueda, T. et al., [1992]Polym. J. 24:1259; Ishihara, K. et al. [1995] Biomaterials 16:873; Campbell et al. [1994] ASAIO J. 40(3):M853; Chen et al. [1996] J. Appl. Polym. Sci. 60:455; and Yamada et al. [1995] JMS Pure Appl. Chem. A32:1723). While these materials have improved stability and promising blood-contacting properties have been reported, a number of limitations exist. In particular, the ability to engineer surface properties on a molecular level, by taking advantage of the principal of self-organization intrinsic to amphiphilic molecules, is lost. In addition, the ability to early incorporate diverse biomolecular functional groups into the membrane-mimetic surface is also lost.
A significant limitation in the widespread use of supported biomembranes is their limited stability for most applications outside of a laboratory environment. In order to generate more robust systems, strategies have been developed to tether membranes to an underlying substrate, such as gold or glass with or without an intervening flexible spacer or polymer cushion (Spinke, J. Y. et al. (1992) Biophys. J. 63:1667-1671; Florin, E.-L. and Gaub, H. E. (1993) Biophys. J 64:375-383; Meuse, C. W. et al, (1998) Biophys J 74:1388-1398; Seitz, M. et al. (1998) Thin Solid Films 329:767-771; Beyer, D. et al. (1996) Thin Solid Films 285, 825-828; Shen, W. W. et al. (2001) Biomacromolecules 2:70-79). While membrane fluidity is critical for many of the functional responses of biological membranes, certain applications lend themselves to compromise in which a substantial increase in membrane stability may be achieved by in situ polymerization of the planar lipid assembly. The capacity to design surfaces with a high degree of molecular control over the assembly of diverse lipid and other membrane-associated constituents is retained. In this regard, applicants and colleagues have previously reported the in situ polymerization of phospholipids on self-assembled monolayers of octadecyl mercaptan bound to gold (Marra, K. G., et al. (1997) Langmuir 13: 5697-5701.), octadecyl trichlorosilane on glass (Marra, K. G., et al. (1997) Macromolecules 30:6483-6488; Orban, J. M. et al. (2000) Macromolecules 33:4205-4212.), and on an amphiphilic polymer cushion (Chon, I. H. et al. (1999) J. Biomat. Sci. Polymer Ed. 10:95-108).
Recently, polyelectrolyte multilayers (PEM) have been studied as bioinert films to reduce cell and protein adhesion (Elbert, D. L. et al. [1999] Langmuir 15:5355-5362) and as coatings to modulate interfacial molecular transport in drug delivery and immunoisolation systems (Moya, S. et al. [2000)] Macromolecules 33:4538-4544; Shi, X. and -Caruso, F. [2001] Langmuir 17:2036-2042). Other potential applications for these materials include their use as ultrafiltration membranes (van Ackern, F. et al. [998] Thin Solid Films 329:762-766) and in the assembly of optoelectronic devices (Cheung, J. H. et al. (1994) Thin Solid Films 244:985).
Membrane-mimetic systems have also had a direct impact on efforts aimed at understanding the mechanisms of blood coagulation at sites of vascular wall injury and on artificial surfaces. One of the most intriguing developments in the past decade has been the recognition that membrane mimetic systems having a phosphorylcholine component limit the induction of surface-associated blood clot formation. This biological property has been attributed to the large amount of surface bound water due to the zwitterion structure of the phosphorylcholine head group. It has also been suggested that specific plasma proteins which inhibit the blood clotting process are selectively adsorbed to this head group (Chapman [1993] Langmuir 9:39).
In a series of investigations using planar membrane models, Thompson and colleagues (Pearce et al. [1993] J. Biol. Chem. 268:22984-22991; and Tendian et al. [1991] Biochemistry 30:10991-10999) have characterized the molecular requirements for prothrombin binding to phospholipid membranes. It has been observed that the phosphorylcholine head group appears to limit the induction of blood clot formation on synthetic surfaces (Ishihara et al. [1994] J. Biomed. Mater. Res. 28:225-232; Hayward et al. [1984] Biomaterials 5:135-142; and Hall et al. [1989] Biomaterials 10[4]:219-224). It has been proposed that this biological property is related to the large amount of water bound to this zwitterionic head group, or conceivably, the selective adsorption to phosphorylcholine of specific plasma protein(s) that inhibit the blood clotting process (Chapman, D. [1993] Langmuir 9:39-45).
Several investigators have described the direct immobilization of thrombomodulin onto polymeric surfaces in order to generate thromboresistant materials for blood contacting applications. Kishida et al. (1994) Biomaterials 15(10):848-852; Kishida et al. (1994) Biomaterials 15(14):1170-1174; and Kishida et al. (1994) ASAIO Journal 40(3):M840-845 have conjugated TM to both aminated and carboxylated surfaces, including poly(vinyl amine) and poly(acrylic acid) surface-grafted polyethylene and a surface-hydrolyzed poly(ether urethaneurea). Vasilets et al. (1997) Biomaterials 18(17):1139-1145, have reported the immobilization of TM onto poly(acrylic acid) surface-grafted PORE. In all cases, the conjugation scheme utilized a carbodiimide based coupling reaction to link TM to the substrate via freely available amino or carboxyl functionalities on the protein surface. In vitro studies demonstrated that both clotting time and protein C activation were enhanced, and this activity appeared to be directly proportional to TM surface density, as determined by a ninhydrin assay. However, the ability to control TM surface concentration was substrate dependent, with reported TM densities ranging between 0.15 and 0.45 μg/cm2, and TM bioactivity was significantly reduced after surface coupling, as evident by protein C activation rates which were increased only 5 to 10-fold as compared with an observed 20,000-fold enhancement when TM is evaluated as a component of either lipid vesicles or the endothelial cell surface. The loss of cofactor activity is believed attributable to the protein immobilization procedure, which is driven by random-site reactions to any accessible functional group on the TM surface, including those within the thrombin binding site. The impact of local flow conditions on the effectiveness of this strategy was not reported.
Although these studies confirm that substrate-bound thrombomodulin has the potential to limit thrombus formation on synthetic surfaces that are otherwise thrombogenic, the observed reduction in thrombomodulin bioactivity emphasizes that thrombomodulin's biological properties are intimately tied to a variety of structural features which can be lost upon direct covalent coupling to a biomaterial surface. For example, thrombomodulin's ability accelerate the thrombin-dependent activation of protein C requires three tandemly repeated EGF-like domains that serve as a thrombin binding site; a serine/threonine-rich 65 A spacer between the EFT-like domains and the transmembrane domain which optimally align thrombin's active site with the critical scissile bond in protein C; and a covalently associated chondroitin supine moiety that increases the affinity of thrombin binding to thrombomodulin by 10- to 20-fold and catalyzes ATIII inactivation of thrombin (Sadler, J. E. [1997] Thromb. Haemostasis 78[1]:392-395; and Esmon, C. T. [1995] FASEB Journal 9[10]:946-955). While some activity is retained even after the solubilization of thrombomodulin with detergents, membrane association significantly accelerates protein C activation by thrombomodulin. This is mediated, in part, by the ability of the membrane to locally concentrate and coordinate the approximate alignment of reacting cofactors and substrates with thrombomodulin (Galvin et al. [1987] J. Biol. Chem. 262[5]:2199-2205). For example, protein C has a C-terminal 4-carboxyglutamic acid (Gla) domain which binds to the cell membrane and presumably increases its local concentration by confining it to the two-dimensional plane of the lipid bilayer (Esmon et al. [1983] J. Biol. Chem. 258:[9]:5548-5553; Mann et al. [1988] Ann. Rev. Biochemistry 57:915-956; Kalafatis et al. [1996] Critical Reviews in Eukaryotic Gene Expression 6[1]:87-101). In addition, the binding of protein C to the plasma membrane may also induce conformational changes that help align the protein C cleavage site with thrombin's proteolytically active domain. Both electrostatic and hydrophobic interactions may be involved in the association of protein C with the cell membrane. In this regard, recent studies suggest that protein C prefers to bind to and function on membranes that contain both phosphatidylcholine and phosphatidylethanolamine lipids. Protein C may also directly interact with fatty acid side chains within the membrane bilayer (Smirnov et al. [1999] Biochemistry 38[12]:3591-3598). It is surprising that the nature of the phospholipid headgroup may contribute to the subsequent catalytic and binding efficiency of activated protein C. For example, Smirnov et al. (1999 supra); and Smirnov et al. (1994) J. Biol. Chem. 269(2):816-819, have demonstrated that both the PE headgroup and phospholipid polyunsaturation contribute to factor Va inactivation by the activated protein C complex Thus, the lipid bilayer serves as an essential ‘cofactor,’ that in concert with thrombomodulin, accelerates protein C activation and subsequently optimizes APC anticoagulant activity.
Atherosclerosis remains a serious source of morbidity and death despite advances in preventive measures and pharmacological therapeutics. Nearly 700,000 vascular surgical procedures are performed annually in the United States along with several hundred thousand peripheral and coronary angioplasties (1). Prosthetic bypass grafts and, more recently, arterial stents and other endovascular prostheses have been utilized in association with these reconstructive procedures. Although large diameter vascular grafts (6 mm internal diameter) have been successfully developed from polymers such as polytetrafluoroethylene (PTFE) and polyethylene terephthalate, the fabrication of a durable small diameter prosthesis (<6 mm internal diameter) remains unsolved. Furthermore, while prosthetic bypass grafting performed in the infra inguinal position with reasonable short-term success, within 5 years 30% to 60% of these grafts will fail (Winger T. M. et al. (1996), “Lipopeptide conjugates: Biomolecular building blocks for receptor activating membrane-mimetic structures. Biomaterials 17:443-449). Likewise, restenosis and/or occlusion occur in as many as 50% of all patients within 6 months of stent placement depending upon the site and the extent of the disease (Winger T. M. et al. [(1997]), “Behavior of lipid-modified peptides in membrane-mimetic monolayers at the air/water interface,” Langmuir 13:3256-3259).
It is recognized that the adverse events leading to the failure of many vascular prostheses are related to maladaptive biological reactions at the blood-material and tissue-material interface. In response to these problems, and particularly thrombosis of the small caliber prosthesis, grafts and stents have been coated with albumin, heparin, or prostacyclin analogues, which inhibit the clotting cascade and platelet reactivity, or with relatively inert materials, such as polyethylene oxide (Marra, K. G. et al. [(1997]), “In-Situ polymerization of phospholipids on an alkylated surface,” Macromolecules 30:6483-6487). An alternate approach has been to design materials which support the in situ regeneration of an endothelial cell lining in order to create a functional arterial substitute with a durable thromboresistant interface (Marra, K. G. et al. [1997], “Stabilized phosphatidylcholine surfaces via in-situ polymerization at a solid-liquid interface,” Polymer Preprints 38(2):682-683; Winger, T. M. and Chaikof, E. L. [1998] “Synthesis and characterization of supported bioactive lipid membranes. In: “Materials Science of the Cell,” Eds. A. Plant and V. Vogel, MRS Publications, Pittsburgh, 1998.) However, strategies based upon the coating or derivation of a prosthetic surface with matrix proteins or integrin-selective peptide sequences that promote endothelial cell growth have been unable to overcome the capacity of these substrates to activate platelets and the coagulation cascade. Thus, in the period prior to complete endothelial regeneration, the surface of a small caliber prosthesis remains at increased risk for thrombus formation.
There is a need in the art for effective antithrombogenic treatments for blood-contacting implants and treatment devices.
Whole organ pancreatic allografts using current immunosuppressive protocols have an expected graft survival as high as 86% at one year and 74% at five years after transplantation. Despite these encouraging results, the risk of major perioperative morbidity, the associated complications of chronic immunosuppressive therapy and the persistent shortage of donor organ tissue remain significant limitations of this approach. As a consequence, in the 1990s pancreas transplantation continues to have a limited role in the management of diabetes. The utilization of xenogeneic organs should provide a solution to the chronic shortage of donor tissue. Nonetheless, the prevention of tissue rejection following cross-species transplantation remains unsolved. It has been postulated that cell based therapy, using xenogeneic islets or insulin producing cell lines in association with an immunoisolation barrier, provides a rational strategy to circumvent the vigorous humoral and cellular response of the host while increasing the supply of non-human donor tissue (Sun, Y. et al. [1996], “Normalization of diabetes in spontaneously diabetic cynomologus monkeys by xenografts of microencapsulated porcine islets without immunosuppression,” J. Clin. Invest. 98:1417; Halle I., et al. [1993] “Protection of islets of Langerhans from antibodies by microencapsulation with alginate-poly-L-lysine membranes,” Transplantation, 44:350-4; Colton, C. and Avgoustiniatos, E. [1991] “Bioengineering in the development of the hybrid artificial pancreas I” Biochem. Eng. 113:152-70; Colton, C. K. [1992], “The engineering of xenogeneic islet transplantation by immunoisolation,” Diab. Nutr. Metabol. 5:145-9).
Cell based therapy, using xenogeneic islets or insulin-producing cell lines in association with an immunoisolation barrier is thus an important step in the successful treatment of insulin-dependent diabetes mellitus (DDM). A critical component of this approach is the maintenance of long-term graft survival by protecting or isolating donor cells from immunological processes in the recipient which lead to islet cell construction. Enhanced control of both transport properties and surface physiochemical characteristics is required for providing the effective immunoisolation barrier crucial to the success of pancreatic islet cell transplantation.
Current approaches for microencapsulation tailor transport properties by controlling the distribution of pore sizes generated by thermodynamically driven physical processes. Typically, semipermeable membranes for cell encapsulation can be formulated by one of three physical processes. The most common methodology is based upon the principle of phase inversion whereby polymer precipitation time, polymer-diluent compatibility, and diluent concentration influence phase separation, and as a consequence, membrane porosity (Crooks, C. A., Douglas, I. A, Broughton, R. L., Sefton, M. V., “Microencapsulation of mammalian cells in a HEMAMMA copolymer: Effects on capsule morphology and permeability,” J. Biomed. Mater. Res. [1990] 24:1241-1262; Sefton, M. V. [1989], Can. I. Chem. Eng. 67:705; Uludag, H. and Sefton, M. V. [1993], “Metabolic activity and proliferation of CEO cells in hydroxymethyl methacrylate methyl methacrylate (KEA-M.A.),” Cell Transplantation 2:175-82; Roberts, I. et al. [1996], “Dopamine secretion by PC12 cells microencapsulated in a hydroxymethyl methacrylate-methyl methacrylate copolymer,” Biomaterials 17:267-75). Using phase inversion techniques, the outer membrane surface morphology can range in pore size from nanometers to microns. As a second approach, barriers can be created by a polyelectrolyte coacervation reaction and molecular weight cutoff (MWCO) modulated by osmotic conditions, diluents, and the molecular weight distribution of the polycationic species (Matthew, H. W. et al [1993] “Complex coacervate microcapsules for mammalian cell culture and artificial organ development,” Biotechnol. Prog. 9:510-519; Yoshioko, T. et al [1990], “Encapsulation of mammalian cell with chitosan-CMC capsule,” Biotechol. Bioeng. 35:66-72; Dautzenberg, H. et al. [1996], Polym. Sci. 101:149). The utilization of multicomponent polycationic polymer blends and the diffusion time of oligocationic species through precast blends of polyanionic polymers have also been shown to be important variables in the control of MWCO when membranes are produced in this fashion. Alginate-calcium chloride systems represent a third technique for generating semipermeable capsules and have been used to produce monodisperse, spherical, transparent beads at a high production rate (Lim, F. and Sun, A. M. [1980], “Microencapsulated islets as a bioartificial endocrine pancreas, Science 210:908; King, G. A. et al [1987], “Alginate-polylysine microcapsules of controlled membrane molecular weight cutoff for mammalian cell culture engineering,” Biotech Progress 3:231-240; Goosen, M. F. A. [1985], “Optimization of microencapsulation parameters: semipermeable microcapsules as a bioartificial pancreas, Biotech Bioeng 27:146-150). As a cell-compatible polysaccharide, alginate is an appealing polymer and, in addition, facilitates cryopreservation of the encapsulated cell (Rajotte, R. et al. [1995], “Adult islet cryopreservation. In: Ricordi, C., ed. “Methods in Cell Transplantation,” Austin: R. Landes, 17-24). Control of transport properties, however, requires post-coating of the alginate with a poly(amino acid), typically, poly-L-lysine or a derivative thereof. In all of these strategies, membrane strength may be increased by altering the composition, structure, and dimensions of the membrane. This often has a secondary effect on diffusive transport. Therefore, in most systems, membrane strength and mass transport properties are interdependent membrane characteristics. It is also significant that the criteria for molecular exclusion is determined, in all of these systems, by the relatively broad distribution of pore sizes and/or dimensional characteristics of non-uniform pores. The inability to precisely control the transport characteristics of currently available encapsulation barriers has lead to their inevitable failure regardless of the chosen membrane formulation.
Recent data suggests that the rejection of pancreatic islet xenografts occurs by an immunological pathway which is distinct from that associated with either the autoimmune destruction of isogeneic islets or rejection of allogeneic grafts. In the latter two cases, islet damage appears to be mediated by a primary “Th1” immune response in which the dominant effector cell is a cytotoxic CD8 T cell (Peterson, I. D., and Haskins, K. [1996], “Transfer of diabetes in the NOD-scid mouse by CD4 T-cell clones: differential requirement for CD8 T-cells,” Diabetes 45:328-36; Haskins, K. and McDuffe, M. [1990], “Acceleration of diabetes in young NOD mice with CD4+ islet-specific T cell clone,” Science 249:1433-6; Jarpe, A. et al. [1990], “Flow cytometric enumeration of mononuclear cell populations infiltrating the islets of Langerhans in prediabetic NOD mice: Development of model of autoimmune insulinitis for Type I diabetes,” Regional Immunology 3:305-17; Miller, B. et al. [1988], “Both the Lyt-2+ and L3T4+ T cell subsets are required for the transfer of diabetes in nonobese diabetic mice I” Immunol. 140:52-8). In contrast, the rejection of islet xenografts is characterized by a ‘Th2’ response in which CD4 helper T cells, but not CD8 cells, play a major role (Weber, C. et al. [1990], “Microencapsulated dog and rat islet xenografts into streptozotocin diabetic and NOD mice,” Horm. Metab. Res. 35:219-226; Akita, K. et al. [1994], “Effect of FK506 and anti-CD4 therapy on fetal pig pancreas xenografts and host lymphoid cells in NOD/Lt, CBA, and BALB/c mice, Cell Transplantation 3:61-73; Gill, R. et al. [1994], “CD4+ T cells are both necessary and sufficient for islet xenograft rejection,” Transplantation Proceedings 26:1203-4; Loudovaris, T. et al. (1992), “The role of T cells in the destruction of xenografts within cell impermeable membranes,” Transplantation Proceedings 24:2938-9; Pierson, R. et al. [1989], “CD-4 positive lymphocytes play a dominant role in murine xenogeneic responses,” Transplantation Proceedings 21:519-21; Parker, W. et al. [1996], “Transplantation of discordant xenografts: a challenge revisited,” Immunology Today 17:373-8). That is, there is considerable evidence that xeno-recognition, unlike allorecognition or autoimmune destruction, primarily occurs via an “indirect” antigen presentation pathway in which host antigen presenting cells (APCs) display peptides scavenged from donor proteins to host helper T cells (Takeuchi, T. et al. [1992], “Heart allografts in murine systems,” Transplantation 53:1281-94; Moses, R. et al. [1990], “Xenogeneic proliferation and lymphokine production are dependent upon CD4+ helper T cells and self antigen-presenting cells in the mouse. I,” Exp. Med. 172:567-75; Lenschow, D. et al. [1992], “Long-term survival of xenogeneic pancreatic islet grafts induced by CTLA4Ig,” Science 257:789-95). Presumably, xenogenic antigens are either released by the occasional broken capsule or diffuse across an intact capsule membrane after having been shed from the cell surface or liberated from necrotic or apoptotic cells. APCs process these antigenic cellular constituents and activate CD4+ T cells which develop into Th2 cells. In turn, Th2 cells stimulate the matron of B cells, which uniquely express the processed foreign peptide, into plasma cells that secrete xenoantigen specific antibodies. Investigators now suspect that it is the generation of immune complexes either by the binding of newly-formed antibodies to xenoantigens or preexisting autoantibodies to islet antigens that leads to inevitable islet cell destruction even in the presence of a barrier which directly excludes their entry. (Weber, C. I. et al. [1998], “Long-term survival of poly-L-lysine-alginate microencapsulated rat, rabbit, and pig islet xenografts in spontaneously diabetic NOD mice,” In: Lanza R, Chick, W., ed. “Handbook of Cell Encapsulation Technology and Therapeutics” [New York: Springer-Verlag]). Antigen-antibody complexes efficiently bind to Fc receptors expressed on the surface of macrophages which leads to their activation and the subsequent secretion of a variety of low molecular weight cytotoxic mediators including cytokines and free radicals, such as interleukin-1 and nitric oxide, respectively (Ke, Y. et al. [1995], “Ovalbumin injected with complete Freund's adjuvant stimulates cytolytic responses,” Eur. J. Immunol. 1995:549-53). It is also likely that activation of the complement cascade by generated antigen-antibody complexes is a significant factor in potentiating islet destruction (Müller-Eberhard, H. I. [1988], “Molecular organization and function of the complement system,” Ann Rev Biochem 57:321-47). The production and association of C3b with circulating immune complexes could enhance the binding of these complexes to macrophages via the cell surface C3b receptor (Krych, M. et al. [1992], Complement receptors,” Curr. Opin. Immunol. 4:8-13). In addition, concomitant release of C3a induces a local neutrophil response with the release of soluble factors, thereby, further potentiating the activation and recruitment of macrophages (Frank, M. [1991], “The role of complement in inflammation and phagocytosis,” Immunol. Today 12:322-6). Notably, at the time of islet rejection an intense pericapsular cellular response is observed which is dominated by the presence of macrophages and B cells (Weber, C. et al. [1994], “NOD mouse peritoneal cellular response to poly-L-lysine-alginate microencapsulated rat islets,” Transplantation Proceedings 26: 1116-1119; Weber, C. I. et al. [1990], “The role of CD4+ helper T cells in destruction of microencapsulated islet xenografts in NOD mice,” Transplantation 49:396-404).
Prior work by inventors hereof includes U.S. patent application Ser. No. 09/149,098 filed Sep. 8, 1998, provisional application No. 60/058,194 filed Sep. 8, 1997, provisional application Nos. 60/091,399 and 60/101,252 filed Jun. 30, 1998 and Sep. 21, 1998, respectively, provisional application No. 60/197,072 filed Apr. 13, 2000, provisional application 60/221,618 filed Jul. 28, 2000, provisional application No. 60/198,792 filed Apr. 20, 2000 and 60/221,828 filed Jul. 28, 2000, PCT application US01/12094 filed Apr. 13, 2001, and PCT application No. US01/12918 filed Apr. 20, 2001, PCT application 97/16080 filed Apr. 11, 1997, application Ser. No. 08/729,928 filed Oct. 15, 1996, Ser. No. 09/342,922 filed Jun. 30, 1999, Ser. No. 09/149,098 filed Sep. 8, 1998, U.S. Pat. No. 6,171,614 issued Jan. 9, 2001, U.S. Pat. No. 6,071,532 issued Jun. 6, 2000, U.S. Pat. No. 5,741,325 issued Apr. 21, 1998, and U.S. Pat. No. 4,906,465 issued Mar. 6, 1990.
All publications referred to herein are incorporated by reference to the extent not inconsistent herewith.